A guide to methods of preserving animal specimens in liquid preservatives. This is adapted from an original set of notes prepared for the Zoo Hons ‘94 field course by Kelvin K. P. Lim & N. Sivasothi http://preserve.sivasothi.com/
- 1 I. OBJECTIVES OF PRESERVATION
- 2 THE PRESERVATIVES
- 3 II. Sample containers
- 4 III. Labelling
- 5 IV. THE ANIMAL GROUPS
I. OBJECTIVES OF PRESERVATION
In this guideline, we are mainly concerned with the taxonomic reasons for preservation. The scientific description of an animal species requires the detailed examination and description of a representative type specimen and a series of specimens which are subsequently deposited, catalogued and maintained in a museum or zoological collection. This remains a reference for other workers to consult in future.
Specimens from any field collection should be deposited in a reference collection in an institutional for the long-term maintenance and access for the future. The animals should therefore be preserved in the best possible condition and where possible, ensure that the natural colour is retained, their external appendages (e.g. fins) are erected and stomach contents intact.
Care should be taken to ensure that specimens are handled carefully. Features important in the taxonomic study of fish, for example, are easily damaged with contact even after preservation. Photography, if required should be conducted on the spot. Do not crowd animals in small containers - this will result in damage to their surfaces or appendages. Do not keep animals for preservation "later" as they may deteriorate quickly and pollute a container leading to a distortion of morphological features and other damage. A well-preserved specimen will generate more accurate information.
In traditional studies, formalin (4% formaldehyde) was used to fix specimens before preserving them long-term in 70% alcohol. However modern studies require samples or whole specimens for molecular studies. In this case the whole specimen or a sample taken from a large specimen should be immersed in 90-97% ethanol. Other media, such as RNAlater, can be used for samples but it does not preserve specimens of some taxa well .
Note: A specimen immersed in a liquid preservative may not be totally infused with the liquid. This may be noticeable when the specimen floats. In the long term, such a specimen will rot, despite being surrounded by a preservative. In large volume animals, injection of the specimen with the preservative is required to reach the inner tissue.
Usage: Formalin should be used only to fix specimens for morphological study. Many groups should NOT be fixed with formalin - cnidarians such as corals, octocorals and sea-pens, echinoderms, crustaceans, and protistans. In most cases once fixed for 24 to 48Hrs depending on the size (longer if large fish), specimens should be transferred to 70% alcohol. Where colour is an important character or needs to be recorded, it is advisable to take photographs of the specimen with a colour swatch.
i. Formalin - 24-48hrs (fix soft tissue)
ii. Water - up to 1 day (depending on size of specimen; to leach out the formalin)
iii. Alcohol - long term storage
These are approximations. The length of time for each step may have to be increased with increasing size of specimens. 10% formalin (= 4% formaldehyde) is usually used to fix specimens. Concentrated formalin should be taken to sea and diluted at sampling sites: Mix one part concentrated formalin to nine parts water. Sea-water does buffer the acidic formalin to some degree but it should not be relied on for long-term storage (beyond several months). Borax is often added to formalin to provide a more consistent buffer allowing longer storage. In bottles meant for killing and fixing, fill about two-thirds the bottle’s volume with 10% formalin.
Warning: Goggles, respirator mask, gloves and protective clothing should be used when handling formaldehyde/formalin. Inhalation of formalin fumes causes extreme discomfort and is harmful. Contact with the fluid causes severe irritation to the skin; contact with sore or raw spots results in extreme pain. Formaldehyde is also said to be a carcinogen. In the event of spillage, always rinse hands, the outside of receptacles and other affected areas thoroughly with water.
Storage: Concentrated or any other forms of formalin should be kept in safe, water-tight, spill-proof bottles, e.g. pep-bottles, and should always be clearly labelled. Plastic bags containing formalin should be leak-proof and maintained in an upright position at all times, and the opening should be rolled or folded down and sealed with tape or clips. Securely cover all receptacles for formalin fumes can be very offensive smelling, and can cause extreme discomfort to the nose and eyes.
2. Alcohol (ethanol)
Usage: Alcohol is usually used for most cnidaria, sponges, arthropods and echinoderms which can be immersed in a solution of 70% if the specimens are being used for morphology but 90-97% pure ethanol if specimens are likely to be used for molecular studies as well. Note that the colour of a specimen is lost almost immediately once immersed in alcohol.
Ethanol usually comes in the 95% concentrated form. For molecular studies use the concentrated form. For long-term preservation, it is usually diluted with fresh water to 70-80% strength. This is the lowest concentration at which preservation will be maintained. During field collections, ensure that solution you use is not diluted by the water which comes with the samples.
Industrial Methylated Spirits (IMS) is also used to preserve specimens at strengths of 70 to 80%. It has added impurities which may affect DNA so should not be used for molecular samples. IMS is, however, cheaper than pure ethanol and does not attract the same custom duties as ethanol. Warning: IMS is poisonous if consumed.
Warning: Alcohol is usually safe to handle, but can cause irritation to the skin in cases of prolonged contact. Always rinse hands thoroughly with water after working with alcohol. Industrial alcohol is toxic and should never be drunk. Receptacles containing alcohol should always be properly and clearly labelled.
Alcohol is highly flammable. Never work with this fluid in the vicinity of open flames. Alcohol is prone to rapid evaporation, and receptacles holding it should be securely covered at all times, and not be opened unnecessarily.
II. Sample containers
There are various types of containers which one can use to fix specimens:
1. Plastic water-tight bottles.
Where possible, the bottle-type containers should be as tall as the length of the largest specimens, such that the specimens can be set straight when the bottle is placed on its side. It can however, be difficult to handle in the field where one has to screw and unscrew the bottle-cap numerous times in a row.
2. Plastic bags.
Not as desirable, but with careful usage, and with knowledge of its limitations, these can be successfully utilised in the absence of plastic containers. Its disadvantage lies mainly in that its leak-proofness is not guaranteed, and also it can easily be punctured by sharp spines. When plastic bags are used, go for those with thicker material. Plastic bags are very useful for fixing the larger specimens, as well as for containing mud and leaf-litter samples.
1. Data labels
Always make sure to insert a data label into the preservation container/bottle for every specimen collected. This should preferably be done immediately before leaving the site for another. A data label is extremely important. It should record the specimen identification if appropriate, cruise number, station number, date, any other supporting data. Specimens without such data are virtually worthless. Data labels should be made from suitable paper so that it does not disintegrate in liquid and should be labelled with waterproof ink or soft pencil. Subsamples of a specimen should also be labelled and note made of the animal and locality data they were take from. Never mix specimens from two different localities in the same bottle. Always use a fresh bottle for each site. Always put a label indicating collection locality. A log of samples should also be made to keep track of specimens and samples collected at each station.
2. Bottle labels
Always label chemical bottles, particularly those with preservatives clearly, so that the contents will not be misused, or worse, drunk accidentally.
IV. THE ANIMAL GROUPS
Whenever possible, fish specimens can be fixed in 10% formalin upon capture, any molecular samples such as from muscle or liver should be taken first and preserved in 97% ethanol or in solutions such as RNALater. When preserving the fish usually try to make sure its finnature is well spread-out, and the body straight and well-stretched. Examination and counting of fin rays and scales would be relatively easy on such well-preserved material.
Specimens are usually left in formalin to be fixed (ie. become thoroughly hardened) for a week or slightly more if larger than 10 cm, before they are soaked for a day or two in clean freshwater before being transferred to 70-75% alcohol for long-term preservation. It is possible to preserve fish specimens for a long time in formalin, but if the preservative is not buffered, the high acidity is likely to render the specimens brittle and transparent.
Large fishes above 10 cm, crustaceans and herptiles should preferably be injected with a small amount of 10% formalin after they have expired in the preservative. Fishes larger than 6 cm should always be injected before they are fixed in formalin. This prevents the innards from rotting, a condition which leads to distortion of the specimens.
When injecting specimens, find a place with running water, wear appropriate protective equipment and clothing. If any spills occur wash hands immediately. Inject 10% formalin into the body cavity of the specimens through the vent until the body bloats (only slightly) and formalin starts to drip from the mouth. For larger fishes an incision could be made on the belly to enable formalin to penetrate the body cavity.
When using bottles, always place fishes with strong spines on their fins into the bottles backwards. This will enable easy extraction of the specimens later, even if the spines become erected, without having to damage the spines.
These are the easiest to process in that they can be killed immediately and stored in alcohol. The larger arthropods (especially those with hard exoskeletons) sometimes need to be injected with preservative to prevent their innards from rotting.
Newly killed molluscs can be fixed in buffered 10% formalin for two or three days, then transferred to 75% alcohol after soaking for a few hours in water.
Different taxa have to be preserved following different protocols. Pennatulacea and Alcyonacea have to be preserved directly in 75% alcohol to prevent dissolving of sclerites that are of taxonomic importance. Scleractinians have to be also preserved in 75% alcohol. Actiniaria, Ceriantharia, Antipatharia and Corallimorpharia are preferably fixed in 10% formol, and small fragment has to be preserved in 96% alcohol for genetic studies. Scyphozoa and Hydrozoa are also preferably fixed in 10% formol.
Preservation of asteroids for subsequent identification and analysis is relatively straightforward. Specimens should be fixed in 95% ethanol, which preserves the specimen in a manner, which makes tissue available for basic DNA extraction. Specimens can be later transferred to 75% ethanol or dried for long-term storage. If histology or other morphological analysis is a high priority for the specimens involved, specimens can be fixed in a buffered formalin solution followed by storage in ethanol or drying. Tube feet from formalin-fixed specimens should be sampled before fixation and preserved in either 95% ethanol or some other DNA-friendly fixative, such as DNA later. If time permits, specimens should be relaxed in freshwater or a magnesium chloride solution prior to fixation.
Ideally, specimens should not be directly dried without fixation, especially if the body wall is heavily thickened, soft or heavily embedded with tissue. However, if adverse circumstances dictate this course of action species with well-developed skeletons can be dry fixed although results may vary. Some individuals will show decay and/or coelomic collapse for example.